Sixty male, 6-week-old C57BL6 mice, weighing 20–22 g were purchased from experimental animal center of Nanjing Medical University of laboratory animal center. All the mice were carefully maintained at room temperature on a 12:12 h light:dark cycle, with free access to chow and water in the cages. This study was approved by the Ethics Committee on Animal Research at the Department of Pediatrics, Huai'an First People's Hospital, Nanjing Medical University, Nanjing, China. The mice were divided into 4 groups: the control group without LPS and ST1926 administration (Con, n=15); LPS-induced group (15 mg/kg, n=15); 10 mg/kg ST1926-treated group after LPS treatment (ST1926/L, n=15); 20 mg/kg ST1926-treated group after LPS (ST1926/H, n=15) (23). After 7 days adaptation, mice from LPS group were treated by intraperitoneal injection with 15 mg/kg body weight LPS for 6 h. The control group was also administered with the same volume of Hanks' buffer. Then, the mice were administered with different concentrations of ST1926 via oral gavage for 2 weeks. The suspension was mixed in high concentration for use immediately after it was ready within 1 min through oral gavage. ST1926, purchased from Sigma-Tau and Biogem (Ariano Irpino, Italy) was dissolved in distilled water. Then, all the mice were sacrificed. Eyeball blood was collected for the next investigation, and the whole lung tissues were carefully harvested on 4°C glacial table.

Mouse lung epithelial cells, MLE-12, and human bronchial epithelial cells (NHBE) were obtained from Shanghai Haoran Biological Technology Co., Ltd. (Shanghai, China) and both cultured in Dulbecco's modified Eagle's medium (DMEM)/F12 containing 1% penicillin/streptomycin and 10% fetal bovine serum (FBS). All cells were cultured in a humidified atmosphere with 5% CO₂ and 95% humidity at 37°C in an incubator. Cells were treated with 100 ng/ml LPS with or without different concentrations (4 µM, ST1926/L; 8 µM, ST1926/H) of ST1926 for 24 h.

Lung epithelial cells (5×10³) were planted into a 96-well plate (Corning Inc., Corning, NY, USA) per well. ST1926, from 0–8 µM, was added to the medium for 24 h. The cells were then incubated at 37°C in an incubator, and the cell viability was detected by the colorimetric MTT assay at 570 nm following the manufacturer's instructions (KeyGen Biotech, Nanjing, China).

Levels of superoxide dismutase (SOD), Catalase (CAT) and malondialdehyde (MDA) in serum and lung tissue samples were determined by commercially available kits (Nanjing Jiancheng Bioengineering Institute, Nanjing, China) according to the manufacturer's instructions.

The serum tumor necrosis factor-α (TNF-α, MTA00B), interleukin-1β (IL-1β, MLB00C) (both from R&D Systems, Minneapolis, MN, USA), IL-8 (BYL02382; Beinglay, Wuhan, China), IL-18 (DY122-05), IL-5 (M5000) (both from R&D Systems), IL-6 (BMS603; Bender MedSystems, Vienna, Austria), IL-12 (PK-EL-62216M; Promocell, Heidelberg, Germany), IL-10 (M1000B; R&D Systems), and IL-17 (BMS6001; Bender MedSystems) were measured according to the manufacturer's protocol using an enzyme-linked immunosorbent assay (ELISA) kit. For the determination of cytokine levels, blood samples from the rats were obtained after sacrifice and stored at −80°C until use.

H₂O₂ levels in lung tissue were detected using a hydrogen peroxide assay kit (catalase assay kit, S0051; Beyotime Institute of Biotechnology, Jiangsu, China) following the manufacturer's instructions. O₂⁻ of lung tissue samples was measured by lucigenin chemiluminescence method. Briefly, lung tissue of mice under different conditions were weighed and homogenized in a homogenization buffer using Hepes and EDTA. After centrifugation, an aliquot of the supernatant was incubated with 5 µM lucigenin in Krebs-Hepes buffer. Light emission was measured with a Tecan Infinite 200. Specificity for O₂⁻ was evaluated by adding 350 U/ml SOD to the incubation medium. Protein concentration was assessed with BCA Protein Quantitative analysis kit (Thermo Fisher Scientific, Waltham, MA, USA).

After treatment, eight mice in each group were sacrificed and BALF was obtained by washing three times with 1 ml of cold sterile phosphate-buffered saline (PBS) through a tracheal cannula. BALF samples were centrifuged at 3,000 rpm for 10 min at 4°C, and the cell pellet was resuspended in PBS for the total cell number assessment with a hemacytometer, and the cytospins were prepared for other differential cell number through staining with the method of Wright-Giemsa staining. Percentages of BALF macrophages, neutrophils and lymphocytes were obtained by counting leukocytes under light microscopy.

Mouse lung tissue lysates were treated with cold ethanol for 1 h at −20°C and then centrifuged at 20,000 × g to remove proteins that can interfere with NO measurements. The potassium iodide-acetic acid reagent was prepared fresh by dissolving 0.05 g of potassium iodide in 7 ml of acetic acid. The KI/AcOH mixture was added into a septum-sealed purge vessel and bubbled with nitrogen gas. The gas stream was connected via a trap containing 1 N NaOH to a Sievers 280i Nitric Oxide Analyzer (GE Analytical Instruments, Boulder, CO, USA). The samples were injected with a syringe through a silicone-Teflon septum. The data were then analyzed via assessing the area under the curve of chemiluminescence signal with the Liquid software (GE Analytical Instruments). The resultant NOx value presents the total nitric oxide and nitrite in pmols/mg protein.

MPO activity in mouse lung tissue was determined using MPO assay kit (BioVision, Milpitas, CA, USA) following the manufacturer's instructions. Briefly, the MPO in the samples catalyzes the production of NaClO from H₂O₂ and NaCl. Next, the NaClO reacts with exogenously added aminophenyl fluorescein to generate fluorescein, which is assessed with a fluorometer at 485 nm excitation and 525 nm emission. The relative fluorescent units of each sample are converted into pmol of fluorescein by a standard curve. The data are reported as pmol fluorescein generated/min/mg of protein extract.

The lung and liver tissue samples were fixed using 10% buffered formalin, imbedded in paraffin and then sliced into 4 µM thick sections. Following the hematoxylin and eosin (H&E) staining, the pathological alterations of the lung tissue samples were observed under a light microscope. Periodic acid-Shiff (PAS) staining was performed using a standard protocol. The number of PAS positive cells was quantified by inspection by a single-blinded investigator. Individual slides were examined independently. Lung tissue samples were IHC stained for the analysis of the lung tissue after different treatments. The sections were stained with TNF-α and IL-1β (Abcam, Cambridge, MA, USA) overnight at 4°C, prior to incubation according to the manufacturer's instructions, followed by horseradish peroxidase conjugated anti-rabbit IgG (Dako, Glostrup, Denmark) as a secondary antibody.

Lung tissue sections and cells were washed twice with PBS and fixed with 3.7% (v/v) formaldehyde in PBS for 15 min. Cells were permeabilised for 5 min with 0.1% Triton X-100, then they blocked with 1% bovine serum albumin for 30 min. For TLR4 and p-NF-κB staining in tissue and cell respectively, 50 µg/ml mouse anti-p-NF-κB and TLR4 antibodies were employed at 4°C overnight, followed by staining with 2 µg/ml Alexa Fluor 488-goat anti-mouse secondary antibodies at room temperature. 4′,6-Diamidino-2-phenylindole (DAPI; Sigma-Aldrich) were used. Images were acquired by confocal laser scanning (Leica TCS SP5; Leica, Heidelberg, Germany) by epifluorescence microscopy (Ningbo Sunny Instruments Co., Ltd., Ningbo, China).

Total RNA was extracted from lung tissue samples and cells by using TRI-reagent (Sigma-Aldrich) following the manufacturer's instructions and treated with deoxyribonuclease I. Then the mRNA was converted into cDNA for real-time PCR analysis. Real-time PCR was carried out (35 cycles of 95°C for 20 sec, 54°C for 30 sec, and 72°C for 30 sec). Fold alterations in mRNA levels of the targeting gene relative to the endogenous cyclophilin control were calculated. Briefly, the cycle threshold (=Ct) values of each target gene were subtracted from the Ct values of the housekeeping gene cyclophilin (ΔCt). Targeting gene ΔΔCt was calculated as ΔCt of target gene minus ΔCt of control. The fold change in mRNA expression was calculated as 2^−ΔΔCt following a previous study. The sequences used in this study are as follows: forward IL-18, (5′-3′) TAA GGA TAC GGA CTA CGG CT and reverse primers, (5′-3′) GTT GGT GGA GGT CTG AGT TTA; forward IL-6, (5′-3′) ACT ACT TATC GTC GAG GTG CTA T and reverse primers, (5′-3′) CGA GCT TGG AGT ACC ATG TTA CTT; forward TNF-α, (5′-3′) ATA GGA ACC AGG GGC AGT T and reverse primers, (5′-3′) CTG CGT TCA GAT GAT TGA TG; forward SOD1, (5′-3′) CTG GCC TGC TCT GCT GCT TGT and reverse primers, (5′-3′) TGG TAG GTG CGG ACT GTG GT; forward SOD2, (5′-3′) CCT TGC GTC ATT CTA TCC A and reverse primers, (5′-3′) GAA ACG CGC CAG AAG GGT TA; forward TLR4, (5′-3′) CTG CTG CCT GCT TGC TGC GT and reverse primers, (5′-3′) GTG GTG TGT GCG GTG TAG AC; forward MyD88, (5′-3′) CAC TTC GCT GTC ATC TCC A and reverse primers, (5′-3′) AGC ACA GAG CGT CAA GAG GT; forward CAT, (5′-3′) CTC TCT GTG ACC GTA ACG CTG GT and reverse primers (5′-3′) TGC GTG TGG TGC GTG ATG GAG ACC; forward haeme oxygenase-1 (HO-1), (5′-3′) CTA GCT TCG TGC ACA GCT GCG T and reverse primers, (5′-3′) GTG ATG ACT GCG CGT GGT GCT AGA AC; forward Nrf2, (5′-3′) CTC TCT GGA CCT TCC GTT GCG CTG T and reverse primers, (5′-3′) GTT GTA GGT CAG TGT TCG AGG ATC; forward TGF-β1, (5′-3′) AGG AAC TAC GCA GTG GGA T and reverse primers, (5′-3′) CGT CTG TCA GAT GAA TGT TG; forward Foxp3, (5′-3′) GCC TCT GGC TTG TCT GCT CTG and reverse primers (5′-3′) GGT ATG GAG TGC GTG ATG TCT G; forward interferon γ (IFNγ), (5′-3′) GCG CTC ATC TCC TCA TAC TAT and reverse primers, (5′-3′) GAA CCA GCC CGG AGA CAG TTG A; forward granzyme B (GzmB), (5′-3′) CCT GAG TCT GAC TGT CGC CAT GT and reverse primers, (5′-3′) GTG TCT GAG TAC TTG GAC GTA; forward Tbx-21, (5′-3′) CAC GTG CTG CTA GCT GGT GCT CAT and reverse primers, (5′-3′) GTC ATG TGC ATG ATA GAT AC; forward glyceraldehyde 3-phosphate dehydrogenase (GAPDH), (5′-3′) CTA AGT CGA ATG CAA ACA GTT CAG and reverse primers, (5′-3′) AAC ATA CCA TCC ACG ACA CGC TC, which was used as the loading control.

In immunoblotting, lung tissue samples and epithelial cells were homogenized into 10% (wt/vol) hypotonic buffer (25 mM Tris-HCl, pH 8.0, 1 mM EDTA, 5 µg/ml leupeptin, 1 mM Pefabloc SC, 50 µg/ml aprotinin, 5 µg/ml soybean trypsin inhibitor, 4 mM benzamidine) to yield a homogenate. Then the final supernatants were obtained by centrifugation at 12,000 rpm for 20 min. Protein concentration was determined by BCA protein assay kit (Thermo Fisher Scientific) with bovine serum albumin as a standard. Then, same amount of total protein were clapped into 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) followed by immunoblotting using the following antibodies: rabbit anti-p-NF-κB (1:1,000), NF-κB (1:1,000) (both from Abcam), SOD1 (1:1,000), SOD2 (1:1,000), TLR4 (1:1,000), MyD88 (1:1,000) (all from Cell Signaling Technology), p38 (1:1,000), p-p38 (1:1,000), extracellular receptor kinase 1/2 (ERK1/2) (1:1,000), p-ERK1/2 (1:1,000), inhibitor-κB kinase (IKKα) (1:1,000), inhibitor-κB kinase-α (IκBα) (1:1,000), TGF-β1 (1:1,000), Foxp3 (1:1,000), IFNγ (1:1,000), CAT (1:1,000) (all from Abcam), HO-1 (1:1,000; Cell Signaling Technology), Nrf2 (1:1,000; Abcam), GzmB (1:1,000; Cell Signaling Technology), Tbx-21 (1:1,000; Abcam), and GAPDH (1:500; Santa Cruz Biotechnology, Inc., Santa Cruz, CA, USA). Western blot bands were observed using GE Healthcare ECL western blotting analysis system and exposed to Kodak X-ray film. Each protein expression level was defined as grey value (version 1.4.2b, Mac OS X, ImageJ; National Institutes of Health, Bethesda, MA, USA) and standardized to housekeeping genes (GAPDH) and expressed as a fold of control.

Data are expressed as the means ± SD. The treated tissues, cells, and corresponding controls were compared using GraphPad PRI SM (version 6.0; GraphPad Software, La Jolla, CA, USA). Significant differences between groups were considered at p<0.05.

As an index reflecting airway inflammation, here the role of ST1926 in inflammation regulation was investigated. First, the bronchoalveolar lavages (BAL) was conducted and explored. As shown in Fig. 1A, LPS treatment resulted in an elevation of the total number of BAL, contributing to an acceleration of neutrophils (Fig. 1B), lymphocytes (Fig. 1C), macrophages (Fig. 1D), and eosinophils (Fig. 1E). Of note, ST1926 significantly reduced the total number of BAL, as well as neutrophils (Fig. 1B), lymphocytes (Fig. 1C), macrophages (Fig. 1D), and eosinophils (Fig. 1E), in a dose-dependent manner. Additionally, in the lung tissue samples with LPS induction, BAL eotaxin was apparently decreased, which was attenuated by ST1926 treatment (Fig. 1F).

LPS induction is well known to cause injury in different organs, including the liver, renal injury and heart, even the lung, as explored previously (24). Here, in order to assess whether ST1926 has potential effects on airway inflammation in children, LPS was used to induce acute lung injury. In this regard, H&E staining was carried out to observe the injuries of lung tissue specimens. Fig. 2A and B shows, that after LPS treatment in mice, the lung tissue sample showed higher inflammation score compared to the Con group, which was comparable. After ST1926 treatment, the inflammatory response was attenuated markedly. In addition, PAS analysis also exhibited that the percent of PAS positive cells was notably upregulated by LPS exposure, whereas reversed in ST1926 administration in a dose-dependent manner (Fig. 2C and D). The data above indicated that ST1926 exhibited protective role against the LPS-induced acute lung injury in mice.

In order to further confirm that ST1926 could attenuate LPS-caused lung injury via inflammation improvement, pro-inflammatory cytokines level in serum were measured. As shown in Fig. 3, we found that pro-inflammatory cytokines of TNF-α (Fig. 3A), IL-6 (Fig. 3B), IL-5 (Fig. 3C), IL-1β (Fig. 3D), IL-18 (Fig. 3E) and IL-17 (Fig. 3F) in serum were considerably stimulated in mice with LPS exposure. In agreement with the results as mentioned above, ST1926 displayed suppressive role in the inflammatory cytokine secretion, suggesting that ST1926, at least partly, improved LPS-induced injury from inflammation inhibition.

TGF-β1 is one of the most studied cytokines that play an essential role in induction and development of acute injury, contributing to the inflammatory cell recruitment (25). GzmB is a serine protease with intracellular and extracellular activities capable of regulating inflammation through cytokine processing and apoptosis of effector cells (26). We tested the hypothesis that TGF-β1 and GzmB will be changed in LPS treatment, and ST1926 may have potential to reverse it. IL-10 secretion is the characteristic activity of cells in regulating inflammatory disease and a potent anti-inflammatory molecule, thereby suppressing the action of many pro-inflammatory and pro-fibrotic molecules. TBX21 is an important transcription factor of adaptive immunity (27–29). As shown in Fig. 4A, we found that TGF-β1 was highly upregulated in LPS treatment, which was reduced due to ST1926 administration from the protein and gene levels, indicating the inflammatory cell accumulation. Foxp3, as another factor of T cell differentiation for lung injury regulation, was also found to be increased in LPS group. Notably, ST1926 significantly decreased its expression in a dose-dependent manner (Fig. 4B). INFγ and GzmB are also reported to be highly expressed in acute lung injury (30). Similarly, INFγ, GzmB and Tbx21 protein and gene levels were apparently enhanced by LPS treatment, which were reduced by ST1926 administration through western blotting and RT-qPCR assays (Fig. 4C–E). In the end, IL-12 and IL-10 protein levels in serum of mice were calculated. IL-12, a proinflammatory cytokine, was upregulated by LPS. ST1926 showed suppressive role in IL-12 expression. In contrast, IL-10, as an important anti-inflammatory cytokine, was found to be reduced by LPS, which was reversed in ST1926 treatment (Fig. 4F and G). Taken together, the data above indicated that ST1926, at least partly had an inhibitory role in regulating inflammatory response.

As mentioned above, inflammation response was observed. Thus, here we attempted to explore how ST1926 altered LPS-induced acute lung injury in mice. First, as shown in Fig. 5A and B, we further confirmed that pro-inflammatory cytokines of TNF-α and IL-1β were significantly stimulated through IHC analysis. ST1926 displayed attenuated role in controlling pro-inflammatory cytokine release. Furthermore, RT-qPCR analysis was carried out to confirm that ST1926 indeed suppressed LPS-caused inflammation response in the lung tissue of mice. As shown in Fig. 5C, TNF-α, IL-18, IL-6 and IL-1β were enhanced for LPS treatment, which was reduced by ST1926 administration in a dose-dependent manner. NF-κB signaling pathway is well known in modulating inflammatory response and controlling pro-inflammatory cytokine secretion, associated with IKKα and IκBα activation (31). As shown in Fig. 5D, western blot analysis indicated that IKKα activation was significantly improved by LPS treatment, resulting in IκBα expression and NF-κB phosphorylation eventually. Interestingly, ST1926 treatment could reverse NF-κB activation through inhibition of IKKα and IκBα expression, impeding inflammatory response. Taken together, ST1926 suppressed LPS-induced acute lung injury by inactivating NF-κB/IκBα.

TLR4/MyD88 signaling pathway has been well explored in inflammation response through NF-κB signaling pathway regulation (32). Thus, here we attempted to explore if TLR4/MyD88 was involved in NF-κB activation. As shown in Fig. 6A and B, TLR4 and MyD88 were found to be upregulated in LPS-treated group, which was in line with NF-κB alteration. ST1926 showed inhibitory role in TLR4 and MyD88 activation on protein levels. Furthermore, RT-qPCR analysis also indicated that LPS induced high gene expression of TLR4 and MyD88, and reversed TLR4 and MtD88 gene levels were observed (Fig. 6C). Immunofluorescent assay was performed to evidence that TLR4 was activated in LPS-treated group, which was reduced for ST1926 administration (Fig. 6D). The data indicated that ST1926 attenuated inflammation response induced by LPS relying on TLR4 signaling pathway suppression.

Oxidative stress is a leading cause in various diseases, including a variety of cancers, and acute liver and renal injury (33). Also, in acute lung injury oxidative stress was also reported (34). Thus, we proposed that ST1926 may perform its role in LPS-induced acute lung injury by inhibiting ROS generation. In order to prove our hypothesis, first in serum and lung tissue samples, SOD and CAT activity was investigated after LPS treatment, activity of SOD and CAT was highly reduced, while MDA was discovered with higher levels in LPS-treated group both in serum and in lung tissue samples (Fig. 7A and B). However, ST1926 administration significantly upregulated SOD and CAT activity, while the MDA levels were reduced. In addition, oxidants of O₂⁻ and H₂O₂ as well as nitrate levels in the LPS-induced lung tissue samples were found to be higher than that in the control group (Fig. 7C). Also, ST1926 showed inhibitory role in expression of these oxidants, which may be another property of ST1926 to attenuate LPS-induced acute lung injury.

In addition, NOx, iNOS and MPO levels were assessed to indicate the extent of lung injury. As shown in Fig. 8A, LPS enhanced NO levels in the tissue samples, which could be reversed by ST1926 administration. Using MPO activity, we found that LPS induced neutrophil infiltration in the lungs of LPS-treated mice, and ST1926 reduced MPO activity (Fig. 8B). MPO activity in pulmonary parenchyma, reflecting the activation of neutrophils, was largely upregulated in acute lung injury condition. NO is produced by the oxidation of l-arginine (l-arg) to l-citrulline and NO, which is catalyzed by NOS. Thus, iNOS is generally considered to be the major contributor to the pathogenesis of acute lung injury (35). In our study, we found that iNOS protein expression levels were highly induced for LPS treatment, which were reversed by ST1926 administration (Fig. 8C). Further, protein levels of MPO, in line with the data above, were promoted in LPS-treated group. The expression levels were downregulated for ST1926 through western blot analysis (Fig. 8C). The data above further indicated that LPS induced acute lung injury due to NOx, MPO and iNOS acceleration, which was in line with previous studies (36,37). ST1926, at least partly, could ameliorate LPS-induced acute lung injury by downregulating the activity of these factors.

The data above indicated that ST1926 suppressed oxidants levels, which is related to upregulated anti-oxidants to improve lung injury. Thus, here we attempted to further explore the possible molecular mechanism regarding ROS. As shown in Fig. 9A, ROS production was highly elevated in LPS-treated group, which was reversed by ST1926 administration. Consistently, anti-oxidants of SOD1 and SOD2 were also upregulated by ST1926 treatment in LPS-induced mice with acute lung injury (Fig. 9B). Finally, p38 and ERK1/2 signaling pathway was investigated to explore whether they were involved in ROS production, mediated by ST1926 in LPS-treated lung tissue samples. As reported previously, p38 and ERK1/2 are of great importance in ROS generation (38). Similarly, in our study, we found that phosphorylated p38 and ERK1/2 were markedly enhanced by LPS treatment, which was reduced by ST1926 administration (Fig. 9C). The data above indicated that ST1926 could reduce ROS production, which was related to p38/ERK1/2 signaling pathway inactivation.

Moreover, this study indicated that in the LPS-induced lung injury, CAT, HO-1 and Nrf2 levels were downregulated in LPS treatment. In contrast, O₂⁻ and H₂O₂ were upregulated. Of note, ST1926 suppressed ROS production by enhancing anti-oxidant expression while reducing oxidants (Fig. 9D). AMP-activated protein kinase (AMPK) is a serine/threonine protein kinase that serves as an energy sensor in the regulation of cellular metabolism. Additionally, AMPK is reported to be closely associated with ROS production (39). Here, in our study, we found that AMPK phosphorylated levels were significantly reduced for LPS, and ST1926 augmented AMPK phosphorylation of Thr172, indicating that AMPK phosphorylation induced by ST1926 may be a key point to modulate ROS generation (Fig. 9E).

Following the results above, we attempted to explore if ST1926 was effective in LPS-induced acute lung injury in vitro by the use of lung epithelial cells. As shown in Fig. 10A, the results indicated that with the increasing of ST1926, even at the highest level, no significant difference was observed on cells. In Fig. 10B, the cell viability showed no significant difference at 8 µM ST1926 treatment for different times until 72 h. In addition, H&E staining suggested that ST1926 at different concentrations exhibited no toxic role in the liver of mice without LPS treatments (Fig. 10C). The data above indicated that ST1926 could ameliorate acute lung injury without any toxicity.

We confirmed that ST1926 showed no toxicity in cells in vitro. Thus, here, we attempted to explore if ST1926 could perform its role in vitro. As shown in Fig. 11A, p-NF-κB was found to be upregulated in LPS-induced lung epithelial cells through immunofluorescence. ST1926 at 4 and 8 µM significantly reduced NF-κB phosphorylated levels. In line with the results in vivo, pro-inflammatory cytokines of TNF-α, IL-1β, IL-18 and IL-6 were apparently upregulated for LPS induction and ST1926 downregulated the inflammatory cytokine release induced by LPS (Fig. 11B). TLR4/MyD88 and IKKα/IκBα signaling pathways were activated by LPS induction, which were inactivated by ST1926 administration (Fig. 11C and D). Additionally, LPS-induced ROS production was decreased in ST1926-treated groups, accompanied by high level of SOD1 and SOD2 (Fig. 11E and F). Further, phosphorylated p38 and ERK1/2 improved by LPS treatment. Significantly, ST1926 administration could inactivate p38 and ERK1/2 activity, ameliorating ROS production (Fig. 11G and H). Following, the cells were exposed to 100 ng/ml LPS for different times, ranging from 0 to 48 h. Next, western blot analysis was used to calculate p38 and ERK1/2 phosphorylation. As shown in Fig. 11I, we found that LPS treatment activated p38 and ERK1/2 levels, and with the increase in treatment time, the phosphorylated form was sustained until 48 h, indicating that ROS may be activated by LPS exposure. As shown in Fig. 11J, we found that LPS significantly upregulated p38 and ERK1/2 phosphorylation, which were reduced for ST1926 in a time-dependent manner. After ST1926 treatment for 12 and 18 h, significant difference was observed in p38 and ERK1/2 activation, respectively. Furthermore, as shown in Fig. 11K, the NF-κB levels in the nucleus were found to be upregulated for LPS treatment, which was in line with p-NF-κB alteration in cytoplasm to enhance pro-inflammatory transcription, contributing to pro-inflammatory cytokines secretion. Of note, ST1926 showed significant role in reducing NF-κB in the nucleus, subsequently downregulating pro-inflammatory cytokine release. In addition, the whole cell IκBα levels were upregulated due to LPS treatment, while p-IκBα was down-regulated. ST1926 could promote IκBα and p-IκBα levels considerably (Fig. 11L). Thus, we supposed that ST1926 could ameliorate inflammation response by reducing NF-κB translocation into the nucleus. The above data indicated that ST1926 indeed attenuated LPS-induced lung injury in vitro via inhibiting inflammation and ROS production.

In this regard, normal human bronchial epithelial cells (NHBE) were included to further explore how ST1926 regulated LPS-induced damage in lung cells. As shown in Fig. 12A, cell viability was investigated with increasing concentrations of ST1926, no significant difference was observed among various groups. Further, 8 µM ST1926 was administered to cells for different times as indicated, ranging from 0 to 90 h. Then, MTT analysis was carried out to investigate the cell viability, which showed no significant change among different groups (Fig. 12B). Inflammatory response and ROS have been revealed to be induced by LPS. The phosphorylated NF-κB and ROS generation was dramatically upregulated by LPS induction (Fig. 12C and D), which were downregulated in ST1926-treated groups, accompanied with the reduced pro-inflammatory cytokines mRNA levels through RT-qPCR analysis (Fig. 12E). Additionally, anti-oxidants of SOD1, SOD2, CAT, HO-1 and Nrf2 were reduced in LPS treatment, contributing to ROS generation, while ST1926 enhanced these anti-oxidants with apparent difference in comparison to the LPS group (Fig. 12F). Next, TLR4/MyD88 signaling pathway was calculated, which was highly activated for LPS, leading to IKKα expression. ST1926 showed inhibitory role in TLR4, MyD88 and IKKα expression induced by LPS (Fig. 12G). NF-κB levels in the nucleus were increased by LPS, while decreased due to ST1926. Accordingly, IκBα and p-IκBα were also augmented in LPS single treatment group, in line with the results in MLE-12 cells mentioned above. ST1926 could reduce IκBα and phosphorylated IκBα expression levels (Fig. 12H). Next, NHBE cells were exposed to LPS for different times as indicated to explore the p-p38 and p-ERK1/2 expression levels. Fig. 12J shows that p38 and ERK1/2 were activated due to LPS treatment, indicating ROS generation. With the increase of treatment time, the phosphorylation of p38 and ERK1/2 was maintained in high levels. Finally, p38 and ERK1/2 protein levels were assessed. As shown in Fig. 12I, phophorylated p38 and ERK1/2 were highly expressed in LPS-treated group, which were reduced by ST1926. In NHBE cells, we found similar results that ST1926 could reduce LPS-induced p38 and ERK1/2 phosphorylation beginning from 12 h with significant difference (Fig. 12K). Thus, the data above indicated that ST1926 could reduce LPS-induced p38 and ERK1/2 phosphorylation from 12 or 18 h in a time-dependent manner.